DNA Protocols & Applications

Considerations for isolation and quantification of both genomic DNA and plasmid DNA

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This section describes considerations for isolation and quantification of both genomic DNA from different sample sources and plasmid DNA. It also deals with common plasmid DNA procedures, including how to make and transform competent cells, how to culture and handle plasmid-containing cells, and commonly used techniques for analysis of genomic DNA.

Genomic DNA

Genomic DNA constitutes the total genetic information of an organism. The genomes of almost all organisms are DNA, the only exceptions being some viruses that have RNA genomes. Genomic DNA molecules are generally large, and in most organisms are organized into DNA–protein complexes called chromosomes. The size, number of chromosomes, and nature of genomic DNA varies between different organisms (see table Sizes and molecular weights of various genomic DNAs). Viral DNA genomes are relatively small and can be single- or double-stranded, linear, or circular. All other organisms have double-stranded DNA genomes. Bacteria have a single, circular chromosome. In eukaryotes, most genomic DNA is located within the nucleus (nuclear DNA) as multiple linear chromosomes of different sizes. Eukaryotic cells additionally contain genomic DNA in the mitochondria and, in plants and lower eukaryotes, the chloroplasts. This DNA is usually a circular molecule and is present as multiple copies within these organelles.

Genomic DNA contains genes, discrete regions that encode a protein or RNA. A gene comprises the coding DNA sequence, as well as the associated regulatory elements that control gene expression. Nuclear eukaryotic genes also contain noncoding regions called introns. The number of genes varies widely between different organisms. Coding DNA represents only a small fraction of eukaryotic genomic DNA: the bulk of the DNA is noncoding, much of which is made up of repetitive sequences. Some noncoding DNA has structural and regulatory functions; however, the function of most of this DNA is largely unknown. The number of copies of each genetic locus present in a cell, or ‘ploidy’, also varies between organisms. The somatic (body) cells of organisms that reproduce sexually are usually diploid, having two sets of homologous chromosomes and hence two copies of each genetic locus, while the germ (reproductive) cells are haploid and have only one copy of each chromosome. Prokaryotic cells are haploid. Some plants are polyploid, for example, modern wheat, which is hexaploid (six copies of each chromosome).

Plasmid DNA

Bacterial plasmids are closed circular molecules of double-stranded DNA that range in size from 1 to >200 kb. They are found in a variety of bacterial species, where they behave as additional genetic units inherited and replicated independently of the bacterial chromosome. However, they rely upon enzymes and proteins provided by the host for their successful transcription and replication.

Plasmids often contain genes that code for enzymes that can be advantageous to the host cell in some circumstances. The encoded enzymes may be involved in resistance to, or production of, antibiotics, resistance to toxins found in the environment (e.g., complex organic compounds), or the production of toxins by the bacteria itself.

Once purified, plasmid DNA can be used in a wide variety of downstream applications such as sequencing, PCR, expression of proteins, transfection, and gene therapy.

DNA can be purified using many different methods and the downstream application determines how pure the DNA should be. In addition to isolation using home-made methods (e.g., CsCl gradients), DNA extraction kits are available from many suppliers. The characteristics of the 3 most common types of DNA extraction kit are shown in the table Characteristics of common DNA extraction kits.

Delivers high-purity nucleic acids for use in most downstream applications

Delivers high-purity nucleic acids for use in most downstream applications

Fast, inexpensive

Fast, inexpensive

No silica-slurry carry over, no alcohol precipitation

Easy to automate; no alcohol precipitation

Anion-exchange methods yield DNA of a purity and biological activity equivalent to at least two rounds of purification in CsCl gradients, in a fraction of the time. Purified nucleic acids are of the highest possible quality and are ideal for sensitive downstream biological applications, such as transfection, microinjection, sequencing, and gene therapy research.

Handling DNA

DNA is a relatively stable molecule. However, introduction of nucleases to DNA solutions should be avoided as these enzymes will degrade DNA. Genomic DNA consists of very large DNA molecules, which are fragile and can break easily. To ensure the integrity of genomic DNA, excessive and rough pipetting and vortexing should be avoided. DNA is subject to acid hydrolysis when stored in water, and should therefore be stored in TE buffer, see table TE buffer, pH 7.4.

Where NX = the number of residues of the respective nucleotide within the oligonucleotide (the MW listed for each nucleotide is the MW of that nucleotide, with associated sodium, incorporated in the oligonucleotide)
For dephosphorylated oligonucleotides: P = –84.0
For phosphorylated oligonucleotides: P = 40.0

The quality of the starting material affects the quality and yield of the isolated DNA. The highest DNA yield and quality is achieved by purifying genomic DNA from freshly harvested tissues and cells. If samples cannot be processed immediately after harvesting, they should be stored under conditions that preserve DNA integrity. In general, genomic DNA yields will decrease if samples, particularly animal samples, are stored at either 2–8°C or –20°C without previous treatment. In addition, repeated freezing and thawing of frozen samples should be avoided as this will lead to genomic DNA of reduced size or to reduced yields of pathogen DNA (e.g., viral DNA). Recommendations for storage of different starting materials are discussed below.

Blood

An anticoagulant should be added to blood samples that will be stored. For example, blood samples treated with heparin or EDTA can be stored at 2–8°C for a few days or at –20°C or –80°C for a few weeks. Alternatively, blood samples can be treated with ACD Solution B (0.48% citric acid, 1.32% sodium citrate, 1.47% glucose; use 1 ml per 6 ml blood) and stored for at least 5 days at 2–8°C or 1 month at –20°C. For long-term storage, blood nuclei can be prepared and stored at –20°C.

Most biological fluids (e.g., plasma, serum, and urine) and stool samples can be stored at 2–8°C for several hours. Freezing at –20°C or –80°C is recommended for long-term storage. Swabs can be stored dry at room temperature.

Formalin fixation and paraffin embedding (FFPE) is another means of sample storage and is particularly relevant for clinical tissue samples. Depending on the tissue type, the speed at which biomolecules are degraded, induced, or modified following harvesting can vary. Therefore, the procedures for tissue removal and fixation should be done as quickly as possible.

Fixation of tissues involves placing specimens in a formalin solution, which can vary in composition (a typical 10% formalin solution may contain 3.7% formaldehyde as well as 1–1.5% methanol). The resulting chemical reaction leads to cross-links between biomolecules, including cross-links between nucleic acids, between proteins, and between nucleic acids and proteins. For optimal results, neutral-buffered formalin solution should be used instead of unbuffered or acidic formalin solutions. Neutral buffer slows down the degradation of formalin, whose degradation products are believed to contribute to impairing nucleic acid quality.

The ratio of formalin to tissue should be at least 10:1 to ensure optimal fixation. This is easy to achieve when working with small tissue specimens, such as needle biopsies. However, when dealing with large tissue samples there may be insufficient formalin for fixation. In this case, sections of the tissue should be cut for formalin fixation. Tissues should be fixed for no more than 24 hours to avoid overfixation.

After fixation in formalin, tissue specimens are embedded in paraffin, a process which consists of several steps. The first step is dehydration, where water is replaced by an alcohol, usually ethanol. This is followed by clearing, where the alcohol is replaced by xylene or a xylene substitute, and by impregnation, where xylene is replaced by paraffin. The final step is embedding, where the entire specimen is surrounded with paraffin. It is important that tissue specimens are fully dehydrated prior to impregnation, as residual water may lead to sample degradation. We recommend always using fresh alcohol and xylene, to avoid any possibility of carryover of water from previous uses. To ensure optimal recovery of usable DNA from FFPE samples, low-melting–temperature paraffin should be used instead. In addition, paraffin containing additives such as beeswax should be avoided, as they may interfere with recovery of biomolecules.

Animal tissue

Freshly harvested tissue can be immediately frozen and stored at –20°C, –80°C, or in liquid nitrogen. Lysed tissue samples can be stored in a suitable lysis buffer for several months at ambient temperature.

Animal and human tissues can also be fixed for storage. We recommend using fixatives such as alcohol and formalin; however, long-term storage of tissues in formalin will result in chemical modification of the DNA. Fixatives that cause cross-linking, such as osmic acid, are not recommended if DNA will be isolated from the tissue. It is also possible to isolate DNA from paraffin-embedded tissue (see Other clinical samples).

Animal, yeast, and bacterial cell cultures

Centrifuge harvested cell cultures, remove the supernatant, and then store the cells at –20°C or –80°C. Alternatively, animal cell nuclei can be prepared and stored at –20°C.

Plant tissue

Fresh leaves and needles from most plant species can be stored for up to 24 hours at 4°C without affecting DNA quality or yield. In general, samples that will be stored for longer than 24 hours should be stored at –80°C. However, some samples (e.g., tree buds) can be stored for several days at 4°C. Tissues stored at 4°C should be kept in a closed container to prevent dehydration. Large samples (e.g., branches) can be stored in a plastic bag containing a wet paper towel.

If it is not practical to store frozen samples, a number of methods are available for drying plant tissue, for example, silica gel, food dehydrators, or lyophilizers (3). To prevent DNA degradation, material should be completely desiccated in less than 24 hours. Dried samples should be kept in the dark at room temperature under desiccating or hermetic conditions for long-term storage. Depending on how the sample was handled, the DNA in herbarium and forensic samples may be degraded. Disrupted plant material can be stored in a suitable lysis buffer for several months at ambient temperature.

Fungal material

Mycelium should be harvested directly from a culture dish or liquid culture. For liquid cultures, the cells should be pelleted by centrifugation and the supernatant removed before DNA isolation or storage. Harvested samples can be either directly frozen or freeze dried, and stored at –80°C.

Complete disruption and lysis of cell walls and plasma membranes of cells and organelles is an absolute requirement for all genomic DNA isolation procedures. Incomplete disruption results in significantly reduced yields.

Disruption methods

Lysis buffer

Disruption generally involves use of a lysis buffer that contains a detergent (for breaking down cellular membranes) and a protease (for digestion of protein cellular components). The choice of protease depends on the lysis buffer used. Some sample types require additional treatment for efficient lysis; this is described in more detail in Special considerations for isolating genomic DNA from different sample sources.

Disruption using rotor–stator homogenizers

Rotor–stator homogenizers thoroughly disrupt animal and plant tissues in 5–90 seconds depending on the toughness of the sample. The rotor turns at very high speed causing the sample to be disrupted by a combination of turbulence and mechanical shearing. Foaming of the sample should be kept to a minimum by using properly sized vessels, by keeping the tip of the homogenizer submerged, and by holding the immersed tip to one side of the tube. Rotor–stator homogenizers are available in different sizes and operate with probes of different sizes. Probes with diameters of 5 mm and 7 mm are suitable for volumes up to 300 µl and can be used for homogenization in microfuge tubes. Probes with a diameter of 10 mm or above require larger tubes.

Disruption using bead mills

In disruption using a bead mill, the sample is agitated at high speed in the presence of beads. Disruption occurs by the shearing and crushing action of the beads as they collide with the cells. Disruption efficiency is influenced by:

Size and composition of beads

Ratio of buffer to beads

Amount of starting material

Speed and configuration of agitator

Disintegration time

The optimal beads to use are 0.1 mm (mean diameter) glass beads for bacteria, 0.5 mm glass beads for yeast and unicellular animal cells, 3–7 mm stainless steel beads for animal tissues, and 3–7 mm stainless steel or tungsten carbide beads for plant and fungal tissues. It is essential that glass beads are pretreated by washing in concentrated nitric acid. Alternatively, use commercially available acid-washed glass beads. All other disruption parameters must be determined empirically for each application.

Disruption using a mortar and pestle

For disruption using a mortar and pestle, freeze the sample immediately in liquid nitrogen and grind to a fine powder under liquid nitrogen. Transfer the suspension (tissue powder and liquid nitrogen) into a liquid-nitrogen–cooled, appropriately sized tube and allow the liquid nitrogen to evaporate without allowing the sample to thaw. Add lysis buffer and continue as quickly as possible with the isolation procedure.

Some sample sources contain substances that can cause problems in DNA isolation and analysis. Special considerations are required when working with these sample sources. In this section, considerations for working with a number of different sources are discussed.

Blood

Human blood samples are routinely collected for clinical analysis. Blood contains a number of enzyme inhibitors that can interfere with downstream DNA analysis. In addition, common anticoagulants such as heparin and EDTA can interfere with downstream assays. DNA isolation from blood requires a method to provide high-quality DNA without contaminants or enzyme inhibitors.

In animals, erythrocytes (red blood cells) from birds, fish, and frogs contain nuclei and hence genomic DNA, while those from mammals do not. Since healthy mammalian blood contains approximately 1000 times more erythrocytes than nuclei-containing leukocytes (white blood cells, comprising lymphocytes, monocytes, and granulocytes) removing the erythrocytes prior to DNA isolation can give higher DNA yields. This can be accomplished by several methods. One is selective lysis of erythrocytes, which are more susceptible than leukocytes to hypotonic shock and burst rapidly in the presence of a hypotonic buffer. Alternatively, Ficoll density-gradient centrifugation can be performed to recover mononuclear cells (lymphocytes and monocytes) and remove erythrocytes. This technique also removes granulocytes. A third method is to prepare a leukocyte-enriched fraction of whole blood, called buffy coat, by centrifuging whole blood at 3300 x g for 10 minutes at room temperature. After centrifugation, three different fractions are distinguishable: the upper clear layer is plasma; the intermediate layer is buffy coat; and the bottom layer contains concentrated erythrocytes.

Blood samples, including those treated to remove erythrocytes, can be efficiently lysed using lysis buffer and protease or proteinase K. Along with the animal’s genomic DNA, viral and bacterial DNA can also be isolated from blood samples.

Other clinical samples

Most biological fluids can be treated in the same way as blood samples for isolation of DNA. Isolation of DNA from stool samples is more difficult, as stool typically contains many compounds that can degrade DNA and inhibit downstream enzymatic reactions.

Animal tissues and cell culture

Animal cell cultures and most animal tissues can be efficiently lysed using lysis buffer and protease or proteinase K. Fresh or frozen samples should be cut into small pieces to aid lysis. Mechanical disruption using a homogenizer or mortar and pestle prior to lysis can reduce lysis time. Skeletal muscle, heart, and skin tissue have an abundance of contractile proteins, connective tissue, and collagen, and care should be taken to ensure complete digestion with protease or proteinase K.

For fixed tissues, the fixative should be removed prior to lysis. Formalin can be removed by washing the tissue in phosphate-buffered saline (PBS). Paraffin should be similarly removed from paraffin-embedded tissues by extraction with xylene followed by washing with ethanol.

Yeast cell cultures

Yeast cell cultures must first be treated with lyticase or zymolase to digest the cell wall. The resulting spheroplasts are collected by centrifugation and then lysed using lysis buffer and proteinase K or protease.

Bacterial DNA can also be isolated from a wide variety of clinical samples. Bacterial cells should be pelleted from biological fluids, and the DNA isolated as for bacterial cell cultures. Swab samples should be pretreated with fungicide before centrifugation of bacterial cells.

DNA viruses

In clinical applications, viral DNA is often (although not always) isolated from cell-free body fluids, where their titer can be very low. Virus particles may need to be concentrated before DNA isolation by ultracentrifugation, ultrafiltration, or precipitation. Addition of carrier DNA may also be necessary during DNA isolation when the expected yield of DNA is low. Integrated viral DNA is prepared using the same procedure as for isolation of genomic DNA from the relevant sample. Bacteriophage, such as M13 and lambda, are isolated from infected bacterial cultures. The bacterial cells must be removed from the culture by centrifugation prior to isolation of viral DNA.

Plants

Isolation of DNA from plant material presents special challenges, and commonly used techniques often require adaptation before they can be used with plant samples. Several plant metabolites have chemical properties similar to those of nucleic acids, and are difficult to remove from DNA preparations. Co-purified metabolites and contaminants introduced by the purification procedure, such as salts or phenol, can inhibit enzymatic reactions or cause variations in UV spectrophotometric measurements and gel migration.

DNA isolation is often improved by using plants grown under conditions that do not induce high levels of plant metabolites. Because of the great variation among plants, it is difficult to make general statements about growth conditions to use. However, as a general guideline, it is recommended to use healthy, young tissues when possible. DNA yields from young tissues are often higher than from old tissue because young tissue generally contains more cells than the same amount of older tissue. Young tissue of the same weight also contains fewer metabolites. In addition, many protocols for “home-made” DNA isolation methods recommend growing plants in darkness for 1–2 days before harvesting to prevent high-level accumulation of plant metabolites.

Good microbiological technique will always ensure the best yield and quality of plasmid DNA. To prepare the perfect bacterial culture for your plasmid prep, follow the steps below.

Prepare a starter culture by inoculating a single colony from a freshly streaked selective plate into 2–10 ml LB (Luria-Bertani) medium containing the appropriate antibiotic. Grow at 37°C for ~8 hours (logarithmic growth phase, see figure Growth of E. coli cultures) with vigorous shaking (~300 rpm). Tip: Do not inoculate directly from glycerol stocks, agar stabs, or plates that have been stored for a long time, as this may lead to loss or mutation of the plasmid. Tip: It is often convenient to grow the starter culture during the day so that the larger culture can be grown overnight for harvesting the following morning.

Dilute the starter culture 1/500 to 1/1000 into a larger volume of selective LB medium, as indicated in the appropriate plasmid purification protocol.
Use a flask of at least 4 times the volume of culture to ensure sufficient aeration.
Do not use a larger culture volume than recommended in the protocol, as this will result in inefficient lysis and reduce the quality of the preparation.

Grow the culture at 37°C with vigorous shaking (~300 rpm) for 12–16 hours (see next section).

Harvest the bacterial culture 12–16 hours after inoculation. This corresponds to the transition from logarithmic into stationary growth phase (see figure Growth curve of E. coli in LB medium), when cell density is high (3–4 x 109 cells per ml) and RNA content of cells is low. Harvesting too early may result in lower than expected yields of plasmid DNA due to a lower cell density. Harvesting too late may result in low plasmid quality and yield due to DNA degradation from over-aging of the culture. Tip: Growth of cultures is dependent on factors such as host strain, plasmid insert and copy number, and culture medium. To determine the optimal harvesting time for a particular system, monitor the cell density and the growth of the culture by measuring the OD600 (see next section).

Harvest the bacterial culture by centrifugation at 6000 x g for 15 min at 4°C. Remove all traces of supernatant by inverting the open centrifuge tube until all of the medium has been drained. The cells are now ready for the lysis procedure, as indicated in the appropriate plasmid purification protocol.
The procedure may be stopped at this point and continued later by freezing the cell pellets obtained by centrifugation. The frozen cell pellets may be stored at –20°C for several weeks.

The E. coli growth curve

The growth curve of an E. coli culture can be divided into several distinct phases. The first, lag phase, occurs directly after dilution of the starter culture into fresh medium. During this phase, cell division is slow as the bacteria adapt to the fresh medium. The bacteria then start to divide more rapidly and the culture enters logarithmic (log) phase (4–5 hours after dilution), during which the number of cells increases exponentially. As the available nutrients in the medium are used up and released metabolites inhibit bacterial growth, the culture becomes saturated and enters stationary phase (~16 hours after dilution), during which cell density remains constant. Eventually the culture enters the phase of decline as cells start to lyse, the number of viable bacteria falls, and DNA becomes partly degraded.

There are different methods for storing E. coli strains depending on the desired storage time. Glycerol stocks and stab cultures enable long-term storage of bacteria, while agar plates can be used for short-term storage. Preparation instructions and useful tips for each of these methods are given below.

Glycerol stocks

E. coli strains can be stored for many years at –70°C in 15% glycerol.

Prepare glycerol stocks of bacteria as follows:

Add 0.15 ml glycerol (100%) to a 2 ml screw-cap vial and sterilize by autoclaving. Tip: Vials of sterilized glycerol can be prepared in batches and stored at room temperature until required.

Add 0.85 ml of a logarithmic-phase E. coli culture to the vial of pre-sterilized glycerol.

Vortex the vial vigorously to ensure even mixing of the bacterial culture and the glycerol.

Freeze in ethanol–dry ice or liquid nitrogen and store at –70°C. Tip: Avoid repeated thawing and re-freezing of glycerol stocks as this can reduce the viability of the bacteria.Tip: For precious strains, storage of 2 stock vials is recommended. Tip: When recovering a stored strain, it is advisable to check the antibiotic markers by streaking the strain onto a selective plate.

Stab cultures

E. coli strains can also be stored for up to 1 year as stabs in soft agar. Stab cultures are used to transport or send bacterial strains to other labs.

Cool the LB agar to below 50°C (when you can hold it comfortably) and add the appropriate antibiotic(s). While the agar is still liquid, add 1 ml agar to a 2 ml screw-cap vial under sterile conditions, then leave to solidify.

Vials of agar can be prepared in batches and stored at room temperature until required.

Using a sterile straight wire, pick a single colony from a freshly streaked plate and stab it deep down into the soft agar several times (see figure Inoculating a stab culture).

Incubate the vial at 37°C for 8–12 h leaving the cap slightly loose.

Seal the vial tightly and store in the dark, preferably at 4°C.

When recovering a stored strain, it is advisable to check the antibiotic markers by streaking the strain onto a selective plate.

Agar plates

Plates of streaked bacteria can be sealed with Parafilm and stored upside-down at 4°C for several weeks. Bacteria should always be streaked onto plates containing the appropriate antibiotic to ensure that selective markers are not lost.

To obtain well-isolated colonies, streak an agar plate as follows:

Flame a wire loop, and cool on a spare sterile agar plate.

Using the wire loop, streak an inoculum of bacteria (from a glycerol stock, stab culture, or single colony on another plate) across one corner of a fresh agar plate, as shown in the figure Streaking bacteria on agar plates.

Flame and cool the wire loop again. Pass it through the first streak and then streak again across a fresh corner of the plate.

Repeat again to form a pattern.

Incubate the plate upside down at 37°C for 12–24 hours until colonies develop.

Generating liquid cultures from bacterial stocks

The figure, Essential steps for storage and handling of E. coli shows the sequence of steps necessary to go from a stored stock of bacteria to a liquid culture for plasmid isolation. Bacterial stocks should always be streaked onto selective plates prior to use, to check that they give rise to healthy colonies carrying the appropriate antibiotic resistance. Stocks can potentially contain mutants arising from the cultures used to prepare them, or can deteriorate during storage.

Inoculate liquid cultures from a healthy, well-isolated colony, picked from a freshly streaked selective plate. This will ensure that cells growing in the culture are all descended from a single founder cell, and have the same genetic makeup.

Tip: Culture volumes >10 ml should not be inoculated directly from a plate, but diluted 1/500 to 1/1000 from a pre-culture of 2–5 ml.

Plasmid specifications

Plasmids vary widely in their copy number (see table Origin of replication and copy numbers of various plasmids and cosmids), depending on the origin of replication they contain (pMB1 or pSC101 for example) which determines whether they are under relaxed or stringent control; as well as the size of the plasmid and its associated insert. Some plasmids, such as the pUC series and derivatives, have mutations which allow them to reach very high copy numbers within the bacterial cell. Plasmids based on pBR322 and many cosmids are generally maintained at lower copy numbers. Very large plasmids are often maintained at very low copy numbers per cell.

Liquid cultures of E. coli can generally be grown in LB (Luria-Bertani) medium. Please note, however, that a number of different LB broths, with different compositions, are commonly used. Different formulations contain different concentrations of NaCl and give rise to varied yields of plasmid DNA. We recommend using the LB composition in the table LB media to obtain highest yields of plasmid DNA.

For preparation of 1 liter of LB medium, add 10 g NaCl, 10 g tryptone, and 5 g yeast extract to 950 ml distilled or deionized water, and shake or stir until dissolved. Adjust the pH to 7.0 with 5 M NaOH. Adjust the volume of the solution to 1 liter with distilled or deionized water. Decant into smaller aliquots and sterilize by autoclaving (see Sterilizing media).

Tip: It is advisable to autoclave liquid medium in several small bottles rather than in one large vessel to avoid possible contamination of an entire batch. After autoclaving, do not use medium for 24 hours to ensure that it is properly sterilized and free of contaminating microorganisms.

Tip: Antibiotics should be added to liquid medium immediately prior to use from stock antibiotic solutions that have been filter-sterilized, distributed into aliquots, and stored in the dark at –20°C (see Antibiotics).

Sterilize liquid or solid media by autoclaving, using a pressure and time period suitable for the type of medium, bottle size, and autoclave type.

Tip: Fill bottles only 3/4 full with medium and loosen the caps before autoclaving to avoid hot medium boiling over. Tighten caps once the media is cool (<40°C) to keep it completely sterile.

Tip: Antibiotics and nutrients such as amino acids are inactivated by the high temperatures of an autoclave. They should be sterilized by filtration through a filter unit with a pore size of 0.2 µm, and added to the cooled, autoclaved medium from properly stored stock solutions.

E. coli strains can generally be streaked and stored on LB plates containing 1.5% agar and the appropriate antibiotic(s).

Preparation: Prepare LB medium according to the composition given in Liquid media. Just before autoclaving, add 15 g agar per liter and mix. After autoclaving, swirl the medium gently to distribute the melted agar evenly throughout the solution. Take care that the hot liquid does not boil over when swirled.

Tip: Cool autoclaved agar medium to below 50°C (when you can hold it comfortably) before adding heat-sensitive antibiotics and nutrients. Mix thoroughly to obtain an even concentration throughout the medium before pouring.

Tip: Pour plates in a laminar-flow hood or, if no hood is available, on a cleaned bench surface next to a Bunsen. Use 30–35 ml medium per standard 90 mm petri dish (~30 plates per liter of medium).

After pouring plates, any air bubbles may be removed by passing the flame of a Bunsen burner briefly over the surface. Do not linger with the flame as this may destroy antibiotics in sections of the plates.

Dry plates either directly after solidification or just before use by removing the lids and standing the plates in a laminar-flow hood for 1 hour. Alternatively, if you do not have access to a hood, plates can be dried with the covers slightly open in a 37°C incubator for 30 min, or left upside down with lids on at room temperature for 2–3 days.

Tip: Store plates inverted at 4°C in a dark room or wrapped in aluminum foil to preserve light-sensitive antibiotics. Do not store for longer than 3 months as antibiotics may degrade.

Bacterial strains carrying plasmids or genes with antibiotic selection markers should always be cultured in liquid or on solid medium containing the selective agent. Lack of antibiotic selection can lead to loss of the plasmid carrying the genetic marker and potentially to selection of faster-growing mutants!

Tip: Prepare stock solutions of antibiotics separately from batches of liquid or solid media, sterilize by filtration, aliquot, and store in the dark at –20°C. Recommended stock and working concentrations for commonly used antibiotics are shown in the table Concentrations of commonly used antibiotics.

Tip: Before adding antibiotics to freshly autoclaved medium, ensure that the medium has cooled to below 50°C.

Effective lysis of bacterial cells is a key step in plasmid isolation as DNA yield and quality depend on the quality of cell lysate used for the purification.

Alkaline lysis

Alkaline lysis is one of the most commonly used methods for lysing bacterial cells prior to plasmid purification (4, 5). Production of alkaline lysates involves four basic steps (see figure The principle of alkaline lysis).

Resuspend harvested bacterial cells in Tris·Cl–EDTA buffer containing RNase A. Tip: Ensure that bacteria are resuspended completely leaving no cell clumps in order to maximize the number of cells exposed to the lysis reagents. Tip: For large-scale purification of low-copy plasmids, for which larger cultures volumes are used, it may be beneficial to increase the lysis buffer volumes in order to increase the efficiency of alkaline lysis and thereby the DNA yield.

Lyse cells using NaOH/SDS. Sodium dodecyl sulfate (SDS) solubilizes the phospholipid and protein components of the cell membrane, leading to lysis and release of the cell contents. NaOH denatures the chromosomal and plasmid DNA, as well as proteins. The presence of RNase A ensures that liberated cellular RNA is digested during lysis. Tip: If after addition of lysis buffer (NaOH/SDS) the solution appears very viscous and is difficult to mix, this indicates excess biomass in the lysate step. This results in insufficient cell lysis and it is recommended to double the amount of lysis and neutralization buffers used. Tip: Avoid vigorous stirring or vortexing of the lysate as this can shear the bacterial chromosome, which will then copurify with the plasmid DNA. The solution should be mixed gently but thoroughly by inverting the lysis vessel 4–6 times. Tip: Do not allow the lysis to proceed for longer than 5 minutes. This is optimal for release of the plasmid DNA, while avoiding irreversible plasmid denaturation.

Neutralize the lysate by adding acidic potassium acetate. Note: The high salt concentration causes potassium dodecyl sulfate (KDS) to precipitate, and denatured proteins, chromosomal DNA, and cellular debris are coprecipitated in insoluble salt-detergent complexes. Plasmid DNA, being circular and covalently closed, renatures correctly and remains in solution. Tip: Precipitation can be enhanced by using chilled neutralization buffer and incubating on ice.

Clear the lysate by either centrifugation or filtration, to precipitate the debris. Note: Purification of plasmid DNA from cleared bacterial lysates was traditionally performed using cesium chloride (CsCl) ultracentrifugation. Today, a variety of commercially available plasmid purification kits offer easy procedures for different throughput requirements and applications.

Other lysis methods

A number of other methods have been described for lysing bacterial cells (1, 6). Some of these methods were developed for other applications and may not be suitable for plasmid DNA preparation.

Boiling lysis: Bacterial cells are treated with lysosome to weaken the cell walls and then lysed by heating in a boiling water bath for ~1 minute.

Lysis with detergent: Bacterial cells are lysed by treatment with and ionic detergent (e.g., SDS) or a nonionic detergent (e.g., Triton X-100).

Cells that have the ability to take up DNA (from a variety of sources) are termed “competent”. Several techniques exist to prepare competent cells and one such technique for preparing competent E. coli is given below.

Note: Cells prepared using this protocol are not suitable for electroporation.

Remove a trace of E. coli cells from the glycerol stock vial with a sterile toothpick or inoculating loop, and streak it out on LB-agar plates containing an appropriate concentration of the relevant selective antibiotic(s) (see Antibiotics). If the host strain has already been cultured and stored at 2–8°C (cultures can be stored at 2–8°C for up to 3 months without any significant loss of viability), streak out bacteria from those stocks.

Positive control to check transformation efficiency

Transform competent cells with 1 ng of a control plasmid containing an antibiotic resistance gene. Plate onto LB-agar plates containing the relevant antibiotic(s). Compare the number of colonies obtained with the control plasmid to the number obtained with the plasmid of interest to compare transformation efficiency.

Negative control to check antibiotic activity

Transform cells with 20 µl of TE. Plate at least 200 µl of the transformation mix on a single LB-agar plate containing the relevant antibiotic(s). An absence of colonies on the plates indicates that the antibiotic is active.

Alcohol precipitation is commonly used for concentrating, desalting, and recovering nucleic acids. Precipitation is mediated by high concentrations of salt and the addition of either isopropanol or ethanol. Since less alcohol is required for isopropanol precipitation, this is the preferred method for precipitating DNA from large volumes. In addition, isopropanol precipitation can be performed at room temperature, which minimizes co precipitation of salt that interferes with downstream applications.

Add 0.6–0.7 volumes of room-temperature isopropanol to the DNA solution and mix well. Tip: Use all solutions at room temperature to minimize co-precipitation of salt. Tip: Do not use polycarbonate tubes for precipitation as polycarbonate is not resistant to isopropanol.

Centrifuge the sample immediately at 10,000–15,000 x g for 15–30 min at 4°C. Tip: Centrifugation should be carried out at 4°C to prevent overheating of the sample. (When precipitating from small volumes, centrifugation may be carried out at room temperature.) Tip: Genomic DNA can alternatively be precipitated by spooling the DNA using a glass rod following addition of isopropanol. The spooled DNA should be transferred immediately to a microfuge tube containing an appropriate buffer and redissolved (see step 9).

Carefully decant the supernatant without disturbing the pellet. Tip: Marking the outside of the tube before centrifugation allows the pellet to be more easily located. Pellets from isopropanol precipitation have a glassy appearance and may be more difficult to see than the fluffy salt-containing pellets resulting from ethanol precipitation.Tip: Care should be taken when removing the supernatant as pellets from isopropanol precipitation are more loosely attached to the side of the tube. Tip: Carefully tip the tube with the pellet on the upper side to avoid dislodging the pellet.Tip: For valuable samples, the supernatant can be retained until recovery of the precipitated DNA has been verified.

Wash the DNA pellet by adding 1–10 ml (depending on the size of the preparation) of room-temperature 70% ethanol. This removes co-precipitated salt and replaces the isopropanol with the more volatile ethanol, making the DNA easier to redissolve.

Centrifuge at 10,000–15,000 x g for 5–15 min at 4°C. Tip: Centrifuge the tube in the same orientation as previously to recover the DNA into a compact pellet.

Carefully decant the supernatant without disturbing the pellet.

Air-dry the pellet for 5–20 min (depending on the size of the pellet). Tip: Do not overdry the pellet (e.g., by using a vacuum evaporator) as this will make DNA, especially high-molecular-weight DNA, difficult to redissolve.

Redissolve the DNA in a suitable buffer. Tip: Choose an appropriate volume of buffer according to the expected DNA yield and the desired final DNA concentration. Tip: Use a buffer with a pH of 7.5–8.0, as DNA does not dissolve easily in acidic buffers. (If using water, check pH.) Tip: Redissolve by rinsing the walls to recover all the DNA, especially if glass tubes have been used. To avoid shearing the DNA, do not pipet or vortex. Tip: High-molecular-weight DNA, such as genomic DNA, should be redissolved very gently to avoid shearing, e.g., at room temperature overnight or at 55°C for 1–2 h with gentle agitation.

Purified DNA should be stored at –20°C or –70°C under slightly basic conditions (e.g., Tris×Cl, pH 8.0 or TE buffer; see tables 1 M Tris×Cl and TE buffer) because acidic conditions can cause hydrolysis of DNA. Avoid repeated freeze-thawing as this will lead to precipitates.

Diluted solutions of nucleic acids (e.g., dilution series used as standards) should be stored in aliquots (in siliconized tubes, where possible) and thawed once only. This avoids adsorption of nucleic acids to the tube walls, which would reduce the concentration of nucleic acids in solution.

Endotoxins, also known as lipopolysaccharides or LPS, are cell membrane components of Gram-negative bacteria (e.g., E. coli). The lipid portion of the outer layer of the outer membrane is completely composed of endotoxin molecules (see figure “Bacterial cell wall”). A single E. coli cell contains about 2 million LPS molecules, each consisting of a hydrophobic lipid A moiety, a complex array of sugar residues and negatively charged phosphate groups (see figure Schematic diagram of the endotoxin molecule). Therefore, each endotoxin molecule possesses hydrophobic, hydrophilic, and charged regions giving it unique features with respect to possible interactions with other molecules. Bacteria shed small amounts of endotoxins into their surroundings while they are actively growing and large amounts when they die. During lysis of bacterial cells for plasmid preparations, endotoxin molecules are released from the outer membrane into the lysate.

Endotoxins significantly reduce transfection efficiencies in endotoxin-sensitive cell lines. Furthermore, endotoxins can influence the uptake of plasmid DNA in transfection experiments by competing with DNA for “free” transfection reagent. Overall, endotoxins represent a non-controllable variable in transfection experiment setup. They are invisible on agarose gels and impossible to detect by optical density and influence the outcome and reproducibility of results and making them difficult to compare and interpret.

Endotoxin contamination of different plasmid preparation methods

The chemical structure and properties of endotoxin molecules and their tendency to form micellar structures lead to copurification of endotoxins with plasmid DNA. For example, in CsCl ultracentrifugation, the CsCl-banded DNA is easily contaminated with endotoxin molecules, which have a similar density in CsCl to plasmid–ethidium bromide complexes.

On size-exclusion resins, the large size of the micellar form of endotoxin causes the molecule to behave like a large DNA molecule; and in anion-exchange chromatography, the negative charges present on the endotoxin molecule can interact with anion-exchange resins, thus leading to copurification of endotoxins with the plasmid DNA.

However, the level of endotoxin contamination found in plasmid DNA is dependent on the purification method used.

How are endotoxins measured?

Historically, endotoxins were measured in a clotting reaction between the endotoxin and a clottable protein in the amoebocytes of Limulus polyphemus, the horseshoe crab.

Today much more sensitive photometric tests (e.g., Kinetic-QCL Test from BioWhittaker, Inc.) are used, which are based on a Limulus amoebocyte lysate (LAL) and a synthetic color-producing substrate. LPS contamination is usually expressed in endotoxin units (EU). Typically, 1 ng LPS corresponds to 1–10 EU.

Influence of endotoxins on biological applications

Endotoxins strongly influence transfection of DNA into primary cells and sensitive cultured cells, and increased endotoxin levels lead to sharply reduced transfection efficiencies. Furthermore, it is extremely important to use endotoxin-free plasmid DNA for gene therapy applications, since endotoxins cause fever, endotoxic shock syndrome, and activation of the complement cascade in animals and humans.

Endotoxins also interfere with in vitro transfection into immune cells such as macrophages and B cells by causing nonspecific activation of immune responses. These responses include the induced synthesis of immune mediators such as IL-1 and prostaglandin. It is important to make sure that plasticware, media, sera, and plasmid DNA are free of LPS contamination to avoid misinterpretation of experimental results.

Endotoxin-free plasticware and glassware

To avoid recontamination of plasmid DNA after initial endotoxin removal, we recommend using only new plasticware which is certified to be pyrogen- or endotoxin-free. Endotoxin-free or pyrogen-free plasticware can be obtained from many different suppliers.

Endotoxins adhere strongly to glassware and are difficult to remove completely during washing. Standard laboratory autoclaving procedures have little or no effect on endotoxin levels. Moreover, if the autoclave has previously been used for bacteria, the glassware will become extensively contaminated with endotoxin molecules. Heating glassware at 180°C overnight is recommended to destroy any attached endotoxin molecules.

It is also important not to recontaminate the purified endotoxin-free DNA by using reagents that are not endotoxin-free.

Reliable measurement of DNA concentration is important for many applications in molecular biology. Spectrophotometry and fluorometry are commonly used to measure both genomic and plasmid DNA concentration. Spectrophotometry can be used to measure microgram quantities of pure DNA samples (i.e., DNA that is not contaminated by proteins, phenol, agarose, or RNA). Fluorometry is more sensitive, allowing measurement of nanogram quantities of DNA, and furthermore, the use of Hoechst 33258 dye allows specific analysis of DNA.

DNA concentration can be determined by measuring the absorbance at 260 nm (A260) in a spectrophotometer using a quartz cuvette. For greatest accuracy, readings should be between 0.1 and 1.0. An absorbance of 1 unit at 260 nm corresponds to 50 µg genomic DNA per ml (A260 =1 for 50 µg/ml; based on a standard 1 cm path length. This relation is valid only for measurements made at neutral pH, therefore, samples should be diluted in a low-salt buffer with neutral pH (e.g., Tris·Cl, pH 7.0). An example of the calculation involved in nucleic acid quantification when using a spectrophotometer (see Spectrophotometric measurement of DNA concentration).

When working with small amounts of DNA, such as purified PCR products or DNA fragments extracted from agarose gels, quantification via agarose gel analysis may be more effective (see Agarose gel).

Tip: If you use more than one cuvette to measure multiple samples, the cuvettes must be matched.

Tip: Spectrophotometric measurements do not differentiate between DNA and RNA, so RNA contamination can lead to overestimation of DNA concentration.

Tip: Phenol has an absorbance maximum of 270–275 nm, which is close to that of DNA. Phenol contamination mimics both higher yields and higher purity, because of an upward shift in the A260 value.

Effects of solvents on spectrophotometric readings

Absorption of nucleic acids depends on the solvent used to dissolve the nucleic acid (7). A260 values are reproducible when using low-salt buffer, but not when using water. This is most likely due to differences in the pH of the water caused by the solvation of CO2 from air. A260/A280 ratios measured in water also give rise to a high variability between readings (see figure Effect of solvent on A260/A280 ratio) and the ratios obtained are typically <1.8, resulting in reduced sensitivity to protein contamination (7). In contrast, A260/A280 ratios measured in a low-salt buffer with slightly alkaline pH are generally reproducible.

Effect of RNA contamination on spectrophotometric readings

Depending on the DNA isolation method used, RNA will be co-purified with genomic DNA. RNA may inhibit some downstream applications, but it will not inhibit PCR. Spectrophotometric measurements do not differentiate between DNA and RNA, so RNA contamination can lead to overestimation of DNA concentration. RNA contamination can sometimes be detected by agarose gel analysis with routine ethidium bromide staining, although not quantified effectively. RNA bands appear faint and smeary and are only detected in amounts ≥25–30 ng (0.5:1 RNA:DNA ratio).

Treatment with RNase A will remove contaminating RNA; this can either be incorporated into the purification procedure or performed after the DNA has been purified. Prior to use, ensure that the RNase A solution has been heat-treated to destroy any contaminating DNase activity. Alternatively, use DNase-free RNase purchased from a reliable supplier.

RNA contamination of plasmid DNA can be a concern depending on the method used for plasmid preparation. Methods using alkaline lysis with phenol extraction cannot separate RNA from plasmid DNA, leading to high levels of RNA contamination. Advanced anion-exchange technology allows isolation of high-molecular-weight genomic DNA that is free of RNA.

Purity of DNA

The ratio of the readings at 260 nm and 280 nm (A260/A280) provides an estimate of DNA purity with respect to contaminants that absorb UV light, such as protein. The A260/A280 ratio is influenced considerably by pH. Since water is not buffered, the pH and the resulting A260/A280 ratio can vary greatly. Lower pH results in a lower A260/A280 ratio and reduced sensitivity to protein contamination (7). For accurate A260/A280 values, we recommend measuring absorbance in a slightly alkaline buffer (e.g., 10 mM Tris·Cl, pH 7.5). Be sure to zero the spectrophotometer with the appropriate buffer.

Pure DNA has an A260/A280 ratio of 1.7–1.9. Scanning the absorbance from 220–320 nm will show whether there are contaminants affecting absorbance at 260 nm. Absorbance scans should show a peak at 260 nm and an overall smooth shape.

Fluorometry allows specific and sensitive measurement of DNA concentration by use of a fluorescent dye; with common dyes including Hoechst dyes and PicoGreen.

Hoechst 33258 has little affinity for RNA, allowing accurate quantification of DNA samples that are contaminated with RNA. It shows increased emission at 458 nm when bound to DNA. DNA standards and samples are mixed with Hoechst 33258 and measured in glass or acrylic cuvettes using a scanning fluorescence spectrophotometer or a dedicated filter fluorometer set at an excitation wavelength of 365 nm and an emission wavelength of 460 nm. The sample measurements are then compared to the standards to determine DNA concentration.

Tip: As Hoechst 33258 preferentially binds AT-rich DNA, use standards with a similar base composition to the sample DNA.

PicoGreen is a highly sensitive measure of dsDNA and can measure as little as 20 pg dsDNA in a 200 µl assay volume. Indeed, DNA concentrations from 500 pg/ml to 500 ng/ml can all be measures using a single dye concentration. The assay is optimized to minimize the fluorescence contributions of RNA and ssDNA, such that dsDNA can be accurately quantified in the presence of equimolar concentrations of ssDNA and RNA with minimal effect on the quantitative results.

Agarose gel analysis enables quick and easy quantification of DNA, especially for small DNA fragments (such as PCR products). As little as 20 ng DNA can be detected by agarose gel electrophoresis with ethidium bromide staining. The DNA sample is run on an agarose gel alongside known amounts of DNA of the same or a similar size. The amount of sample DNA loaded can be estimated by comparison of the band intensity with the standards either visually (see figure Agarose gel analysis of plasmid DNA) or using a scanner or imaging system. Be sure to use standards of roughly the same size as the fragment of interest to ensure reliable estimation of the DNA quantity, since large fragments interchelate more dye than small fragments and give a greater band intensity.

More precise agarose gel quantification can be achieved by densitometric measurement of band intensity and comparison with a standard curve generated using DNA of a known concentration. In most experiments the effective range for comparative densitometric quantification is between 20 and 100 ng.

Tip: The amount of DNA used for densitometric quantification should fall within the linear range of the standard curve.

Principle of restriction digestion

Many applications require conversion of genomic DNA into conveniently sized fragments by restriction endonuclease digestion. This yields DNA fragments of a convenient size for downstream manipulations. Restriction endonucleases are bacterial enzymes that bind and cleave DNA at specific target sequences. Type II restriction enzymes are the most widely used in molecular biology applications. They bind DNA at a specific recognition site, consisting of a short palindromic sequence, and cleave within this site, e.g., AGCT (for AluI), GAATTC (for EcoRI), and so on. Isoschizomers are different enzymes that share the same specificity, and in some cases, the same cleavage pattern.

Tip: Isoschizomers may have slightly different properties that can be very useful. For example, the enzymes MboI and Sau3A have the same sequence specificities, but MboI does not cleave methylated DNA, while Sau3A does. Sau3A can therefore be used instead of MboI where necessary.

Selecting suitable restriction endonucleases

The following factors need to be considered when choosing suitable restriction enzymes:

Fragment size

Methylation sensitivity

Blunt-ended/sticky-ended fragments

Compatibility of reaction conditions (where more than one enzyme is used)

Fragment size

Restriction enzymes with shorter recognition sequences cut more frequently than those with longer recognition sequences. For example, a 4 base pair (bp) cutter will cleave, on average, every 44 (256) bases, while a 6 bp cutter cleaves every 46 (4096) bases.

Tip: Use 6 bp cutters for mapping genomic DNA or YACs, BACs, or P1s, as these give fragments in a suitable size range for cloning.

Methylation

Many organisms have enzymes called methylases that methylate DNA at specific sequences. Not all restriction enzymes can cleave their recognition site when it is methylated. Therefore the choice of restriction enzyme is affected by its sensitivity to methylation. In addition, methylation patterns differ in different species, also affecting the choice of restriction enzyme.

The CpG dinucleotide occurs about 5 times less frequently in mammalian DNA than would be expected by chance, and most restriction enzymes with a CpG dinucleotide in their recognition site do not cleave if the cytosine is methylated. Therefore many enzymes with CpG in their recognition site, such as EagI, NotI, and SalI, cleave mammalian DNA only rarely.

Drosophila, Caenorhabditis, and some other species do not possess methylated DNA, and have a higher proportion of CpG dinucleotides than mammalian species. Rare-cutter enzymes therefore cleave more frequently in these species.

Plant DNA is highly methylated, so for successful mapping in plants, choose enzymes that either do not contain a CpG dinucleotide in their recognition site (e.g., DraI or SspI) or that can cleave methylated CpG dinucleotides (e.g., BamHI, KpnI, or TaqI).

Tip: Methylation patterns differ between bacteria and eukaryotes, so restriction patterns of cloned and uncloned DNA may differ.

Blunt-ended/sticky-ended fragments

Some restriction enzymes cut in the middle of their recognition site, creating blunt-ended DNA fragments. However, the majority of enzymes make cuts staggered on each strand, resulting in a few base pairs of single-stranded DNA at each end of the fragment, known as “sticky” ends. Some enzymes create 5' overhangs and others create 3' overhangs. The type of digestion affects the ease of downstream cloning:

Sticky-ended fragments can be easily ligated to other sticky-ended fragments with compatible single-stranded overhangs, resulting in efficient cloning.

Blunt-ended fragments usually ligate much less efficiently, making cloning more difficult. However, any blunt-ended fragment can be ligated to any other, so blunt-cutting enzymes are used when compatible sticky-ended fragments cannot be generated – for example, if the polylinker site of a vector does not contain an enzyme site compatible with the fragment being cloned.

Compatibility of reaction conditions

If a DNA fragment is to be cut with more than one enzyme, both enzymes can be added to the reaction simultaneously provided that they are both active in the same buffer and at the same temperature. If the enzymes do not have compatible reaction conditions, it is necessary to carry out one digestion, purify the reaction products, and then perform the second digestion.

Restriction digest components

Water

DNA

Buffer

Enzyme

The amount of DNA digested depends on the downstream application and the genome size of the organism being analyzed. We recommend using a minimum of 10 µg DNA per reaction for Southern blotting of mammalian and plant genomic DNA. For mapping of cloned DNA, 0.2–1 µg DNA per reaction is adequate.

Tip: DNA should be free from contaminants such as phenol, chloroform, ethanol, detergents, or salt, as these may interfere with restriction endonuclease activity.

One unit of restriction endonuclease completely digests 1 µg of substrate DNA in 1 hour. However, supercoiled plasmid DNA generally requires more than 1 unit/µg to be digested completely. Most researchers add a 10-fold excess of enzyme to their reactions in order to ensure complete cleavage.

Tip: Ensure that the restriction enzyme does not exceed more than 10% of the total reaction volume; otherwise the glycerol in which the enzyme is supplied may inhibit digestion.

Reaction volume

Most digests are carried out in a volume between 10 and 50 µl. (Reaction volumes smaller than 10 µl are susceptible to pipetting errors, and are not recommended.)

Setting up a restriction digestion

Pipet reaction components into a tube and mix well by pipetting. Tip: Thorough mixing is extremely important. Tip: The enzyme should be kept on ice and added last. Tip: When setting up large numbers of digests, make a reaction master mix consisting of water, buffer, and enzyme, and aliquot this into tubes containing the DNA to be digested.

Centrifuge the tube briefly to collect the liquid at the bottom.

Incubate the digest in a water bath or heating block, usually for 1–4 h at 37°C. However, some restriction enzymes require higher (e.g., 50–65°C) while others require lower (e.g., 25°C) incubation temperatures.

For some downstream applications it is necessary to heat-inactivate the enzyme after digestion. Heating the reaction to 65°C for 20 min after digestion inactivates the majority of enzymes that have optimal incubation temperature of 37°C. Note that some restriction enzymes are not fully inactivated by heat treatment.

In order to construct new DNA molecules, DNA must first be digested using restriction endonucleases (see Restriction endonuclease digestion of DNA). The individual components of the desired DNA molecule are purified and then combined and treated with DNA ligase. The products of the ligation mixture are introduced into competent E. coli cells and transformants are identified by appropriate genetic selection. Appropriate control ligations should also be performed (see Preparation of competent E. coli and Transformation of competent E. coli).

Removal of 5' phosphates from linearized vector DNA can help prevent vector self-ligation and improve ligation efficiency. To remove 5' phosphates from DNA, add calf intestinal phosphate (CIP) buffer and 1 U CIP and incubate for 30–60 minutes at 37°C. Once the reaction is complete, inactivate CIP by heating to 75°C for 15 minutes.

Incubate for 1–24 h at 15°C. Tip: Simple ligations with two fragments having 4 bp 3' or 5' overhanging ends require much less ligase than more complex ligations or blunt-end ligations. The quality of the DNA will also affect the amount of ligase needed. Tip: Ligation of sticky-ends is usually carried out at 12–15°C to maintain a balance between annealing of the ends and the activity of the enzyme. Higher temperatures make annealing of the ends difficult, while lower temperatures diminish ligase activity. Tip: Blunt-end ligations are usually performed at room temperature since annealing is not a factor, though the enzyme is unstable above 30°C. Blunt-end ligations require about 10–100-times more enzyme than sticky-end ligations in order to achieve an equal efficiency.

From individual E. coli transformants, purify plasmid or phage DNAs by miniprep procedure and determine their structures by restriction mapping. Tip: It is recommended to include two controls in every transformation experiment: A “mock” transformation without DNA and a transformation with a known amount of closed circular plasmid DNA.

Controls are essential if things go wrong. For example, colonies on plates that receive mock-transformed bacteria may indicate that the medium lacks the correct antibiotic. An absence of colonies on plates receiving bacteria transformed with plasmids under construction can only be interpreted if a positive control using a standard DNA has been included. See Bacterial cultivation media and antibiotics for further information on transformation controls.

Gels allow separation and identification of nucleic acids based on charge migration. Migration of nucleic acid molecules in an electric field is determined by size and conformation, allowing nucleic fragments of different sizes to be separated. However, the relationship between the fragment size and rate of migration is non-linear, since larger fragments have greater frictional drag and are less efficient at migrating through the polymer.

Agarose gel analysis is the most commonly used method for analyzing DNA fragments between 0.1 and 25 kb, while pulse-field gel electrophoresis enables analysis of DNA fragments up to 10,000 kb. This section provides useful hints for effective gel analysis of DNA.

Agarose concentration

The concentration of agarose used for the gel depends primarily on the size of the DNA fragments to be analyzed. Low agarose concentrations are used to separate large DNA fragments, while high agarose concentrations allow resolution of small DNA fragments (see table Concentration of agarose used for separating fragments of different sizes). Most gels are run using standard agarose, although some special types of agarose are available for particular applications. For example, low-melt agarose allows in situ enzymatic reactions and can therefore be used for preparative gels. Genomic DNA can be isolated directly from cells immobilized in low-melt agarose gels (see reference 6 for more information).

Tip: Use ultrapure-quality agarose since impurities such as polysaccharides, salts, and proteins can affect the migration of DNA. Agarose quality is particularly important when running high-percentage agarose gels.

Adapted from references 1 and 6.
* Most gels are run using standard agarose, although some special types of agarose are available for particular applications, and for very high or low agarose concentrations. For example, low-melt agarose allows in situ enzymatic reactions and can be used for preparative gels.

Electrophoresis buffers

The most commonly used buffers for agarose gel electrophoresis are TBE (Tris·borate–EDTA) and TAE (Tris·acetate–EDTA) (see tables TAE, TBE, and Gel loading buffer). Although more frequently used, TAE has a lower buffering capacity than TBE and is more easily exhausted during extended electrophoresis. TBE gives better resolution and sharper bands, and is particularly recommended for analyzing fragments <1 kb.

The drawback of TBE is that the borate ions in the buffer form complexes with the cis-diol groups of sugar monomers and polymers, making it difficult to extract DNA fragments from TBE gels using traditional methods.

* 15% Ficoll (Type 400) or 30% glycerol can be used instead of sucrose.

Pouring the gel

Prepare enough 1x electrophoresis buffer both to pour the gel and fill the electrophoresis tank.

Add an appropriate amount of agarose (depending on the concentration required) to an appropriate volume of electrophoresis buffer (depending on the type of electrophoresis apparatus being used) in a flask or bottle. Tip: The vessel should not be more than half full. Cover the vessel to minimize evaporation. Tip: Always use the same batch of buffer to prepare the agarose as to run the gel since small differences in ionic strength can affect migration of DNA.

Heat the slurry in a microwave or boiling water bath, swirling the vessel occasionally, until the agarose is dissolved. Tip: Ensure that the lid of the flask is loose to avoid build-up of pressure. Be careful not to let the agarose solution boil over as it becomes super-heated.Tip: If the volume of liquid reduces considerably during heating due to evaporation, make up to the original volume with distilled water. This will ensure that the agarose concentration is correct and that the gel and the electrophoresis buffer have the same buffer composition.

Cool the agarose to 55–60°C.

Pour the agarose solution onto the gel tray to a thickness of 3–5 mm. Insert the comb either before or immediately after pouring the gel. Leave the gel to set (30–40 min). Tip: Ensure that there is enough space between the bottom of the comb and the glass plate (0.5–1.0 mm) to allow proper formation of the wells and avoid sample leakage. Tip: Make sure that there are no air bubbles in the gel or trapped between the wells.

Carefully remove the comb and adhesive tape, if used, from the gel. Fill the tank containing the gel with electrophoresis buffer. Tip: Add enough buffer to cover the gel with a depth of approximately 1 mm liquid above the surface of the gel. If too much buffer is used the electric current will flow through the buffer instead of the gel.

Agarose gel electrophoresis allows analysis of DNA fragments between 0.1 and 25 kb (e.g., genomic DNA digested with a frequently cutting restriction endonuclease), while pulse-field gel electrophoresis enables analysis of DNA fragments up to 10,000 kb (e.g., undigested genomic DNA or genomic DNA digested with rare cutting restriction endonucleases). The amount of genomic DNA loaded onto a gel depends on the application, but in general, loading of too much DNA should be avoided as this will result in smearing of the DNA bands on the gel.

Gel loading buffer (see table Gel loading buffer) must be added to the samples before loading and serves three main purposes:

To increase the density of the samples to ensure that they sink into the wells on loading.

To add color to the samples through use of dyes such as bromophenol blue or xylene cyanol, facilitating loading.

To allow tracking of the electrophoresis due to co-migration of the dyes with DNA fragments of a specific size.

Preparation of samples

Add 1 volume of gel loading buffer to 6 volumes DNA sample and mix.
Samples should always be mixed with gel loading buffer prior to loading on a gel. Tip: Do not use sample volumes close to the capacity of the wells, as samples may spill over into adjacent wells during loading. Tip: Be sure that all samples have the same buffer composition. High salt concentrations, for example in some restriction buffers, will retard the migration of the DNA fragments.
Ensure that no ethanol is present in the samples, as this will cause samples to float out of the wells on loading.

Agarose gel electrophoresis

Apply samples in gel loading buffer to the wells of the gel.
Prior to sample loading, remove air bubbles from the wells by rinsing them with electrophoresis buffer. Tip: Make sure that the entire gel is submerged in the electrophoresis buffer. Tip: To load samples, insert the pipet tip deep into the well and expel the liquid slowly. Take care not to break the agarose with the pipet tip. Tip: Once samples are loaded, do not move the gel tray/tank as this may cause samples to float out of the wells. Tip: Be sure to always include at least one lane of appropriate molecular-weight markers.

Connect the electrodes so that the DNA will migrate towards the anode (positive electrode). Tip: Electrophoresis apparatus should always be covered to protect against electric shocks.

Turn on the power supply and run the gel at 1–10 V/cm until the dyes have migrated an appropriate distance. This will depend on the size of DNA being analyzed, the concentration of agarose in the gel, and the separation required. Tip: Avoid use of very high voltages which can cause trailing and smearing of DNA bands in the gel, particularly with high-molecular-weight DNA. Tip: Monitor the temperature of the buffer periodically during the run. If the buffer becomes overheated, reduce the voltage. Tip: Melting of an agarose gel during the electrophoresis is a sign that the buffer may have been incorrectly prepared or has become exhausted during the run. Tip: For very long runs, e.g., overnight runs, use a pump to recycle the buffer.

Pulse-field gel electrophoresis

Apply samples in gel loading buffer to the wells of the gel. Tip: Pulse-field gel electrophoresis uses high voltages, so TBE buffer, which has greater buffering capacity than TAE buffer, should be used. Tip: Prior to sample loading, remove air bubbles from the wells by rinsing them with electrophoresis buffer. Tip: Make sure that the entire gel is submerged in the running buffer. Tip: To load samples, insert the pipet tip deep into the well and expel the liquid slowly. Take care not to break the agarose with the pipet tip. Tip: Once samples are loaded, do not move the gel tray/tank as this may cause samples to float out of the wells. Tip: Be sure to always include at least one lane of appropriate molecular-weight markers.

Connect the electrodes so that the DNA will migrate towards the anode (positive electrode). Tip: Electrophoresis apparatus should always be covered to protect against electric shocks.

Turn on the power supply and run the gel at 170 V with a switch interval of 5–40 s until the dyes have migrated an appropriate distance. This will depend on the size of DNA being analyzed, the concentration of agarose in the gel, and the separation required. Tip: Monitor the temperature of the buffer periodically during the run. If the buffer becomes heated, reduce the voltage. Tip: Melting of an agarose gel during the electrophoresis is a sign that the buffer may have been incorrectly prepared or has become exhausted during the run. Tip: For very long runs, e.g., overnight runs, use a pump to recycle the buffer.

Staining

To allow visualization of the DNA samples, agarose gels are stained with an appropriate dye. The most commonly used dye is the intercalating fluorescent dye ethidium bromide, which can be added either before or after the electrophoresis (see table Comparison of ethidium bromide staining methods). Alternatives include recently introduced commercial dyes such as SYBR Green.

Tip: Stock solutions of ethidium bromide (generally 10 mg/ml) should be stored at 4°C in a dark bottle or bottle wrapped in aluminum foil.

Addition of ethidium bromide prior to electrophoresis — add ethidium bromide at a concentration of 0.5 µg/ml to the melted and subsequently cooled agarose, that is, just before pouring the gel.

Mix the agarose–ethidium bromide solution well to avoid localized staining.

Addition of ethidium bromide after electrophoresis — soak the gel in a 0.5 µg/ml solution of ethidium bromide (in water or electrophoresis buffer) for 30–40 minutes.

Tip: Rinse the gel with buffer or water before examining it to remove excess ethidium bromide.

Tip: Staining buffer can be saved and re-used.

Note: Ethidium bromide is a powerful mutagen and is very toxic. Wear gloves and take appropriate safety precautions when handling. Use of nitrile gloves is recommended as latex gloves may not provide full protection. After use, ethidium bromide solutions should be decontaminated as described in commonly used manuals (1, 6).

Visualization

Ethidium bromide–DNA complexes display increased fluorescence compared to the dye in solution. This means that illumination of a stained gel under UV light (254–366 nm) allows bands of DNA to be visualized against a background of unbound dye. The gel image can be recorded by taking a Polaroid photograph or using a gel documentation system.

Tip: UV light can damage the eyes and skin. Always wear suitable eye and face protection when working with a UV light source.

Tip: UV light damages DNA. If DNA fragments are to be extracted from the gel, use a lower intensity UV source if possible and minimize exposure of the DNA to the UV light.

Southern blotting is a widely used technique that allows analysis of specific DNA sequences. DNA is usually first converted into conveniently sized fragments by restriction digestion. The DNA is next run through an agarose gel (6). Southern blotting (named after its inventor, E.M. Southern) refers to the transfer of the DNA to a nylon or nitrocellulose membrane by capillary transfer. The DNA of interest can be identified by hybridization to radioactive or chemiluminescent probes and visualized by autoradiography or staining.

Many variations on the Southern blotting procedure exist. A standard protocol is described here together with recipes for buffers and solutions.

Preparation of gels for Southern blotting

Fragmentation of large DNA molecules (optional)

DNA fragments longer than 10 kb do not transfer to blotting membranes efficiently. In order to facilitate their transfer, these fragments are reduced in size, either by acid depurination or by UV irradiation.

Acid depurination — immediately after gel electrophoresis, place the gel in a solution of 0.2 M HCl, so that it is completely covered. Agitate gently for 10 minutes. During this period the color of the bromophenol blue in the samples will change from blue to yellow, indicating that the gel has been completely saturated with the acid. Rinse the gel briefly in distilled water.

Tip: The depurination step should not last too long, since very short fragments attach less firmly to the membrane.

Tip: Depurinated gels may yield “fuzzy” bands on the final autoradiograph, presumably because of increased diffusion of the DNA during transfer. Depurination is therefore recommended only when fragments larger than 10 kb are to be transferred.

UV irradiation — expose the gel to UV light at a wavelength of 254 nm from a source operating at 30 W, for 30–60 seconds.

Denaturation

Double-stranded DNA must be denatured in order to create suitable hybridization targets. Completely cover the gel with denaturation buffer (see table Denaturation buffer) and incubate for 30 minutes with gentle shaking. If acid depurination was used to denature the DNA, the bromophenol blue will return to its original color during this incubation.

Assembling the blotting apparatus

Place a support larger than the gel in a tray containing 10x SSC (see table 20x SSC), and cover the support with a glass or Plexiglas plate (see figure Southern blot setup).

Cut two lengths of Whatman 3MM paper wider than the gel, long enough to fit under the gel and reach to the bottom of the dish on either side. Wet the sheets briefly in 10x SSC, and place them on the glass plate. Remove any air bubbles between the paper and the support by rolling a pipet several times back and forth over the surface.

Cut one sheet of blotting membrane and two sheets of Whatman 3MM paper about 1 mm larger than the gel on each side. Tip: Always wear gloves when working with blotting membranes. Handle membranes carefully by the edges or using clean blunt-ended forceps.

Place the prepared gel upside-down on the platform. Remove any air bubbles trapped between the gel and the platform by rolling a pipet several times back and forth over the gel.

Surround the gel with plastic wrap. This ensures that the 10x SSC moves only through the gel and not around it.

Place the precut blotting membrane on top of the gel so that it covers the entire surface. Do not move the blotting membrane once it has been placed on the gel. Remove any air bubbles between the paper and the support as described in step 4.

Briefly wet the two precut sheets of Whatman 3MM paper in 10x SSC, and place them on top of the nylon membrane. Again, remove any trapped air bubbles as described in step 4.

Place a 15–20 cm stack of dry paper towels on top of the filter paper. Tip: Make sure that the plastic wrap surrounding the gel prevents contact of the paper towels with the 10x SSC and the wet filter paper under the gel. Ensure that the towels do not droop over since they can cause liquid to flow around the gel instead of through it.

Place a second glass or Plexiglas plate on top of the paper towels. Place the weight on top of the plate.Let the transfer proceed for 12–18 h. Tip: Transfer efficiency is improved by removing the wet paper towels and replacing them with dry ones at least once during the transfer.

Fixing the DNA to the blot

After the transfer is complete, remove the weight, paper towels, and the two sheets of filter paper. Turn over the gel and the blotting membrane together, and lay them, gel-side up, on a sheet of dry filter paper. Mark the positions of the gel lanes on the membrane using a ballpoint pen or a soft-lead pencil. Peel the gel from the membrane. If desired, keep the gel to assess the efficiency of DNA transfer, otherwise discard.
Before removing the gel from the blotting membrane, ensure that the gel lanes are marked so that they can be later identified.
In order to assess the efficiency of DNA transfer, stain the gel with ethidium bromide after blotting to see how much DNA remains.

Fix the DNA to the blot, either by baking (see step 3) or by UV-crosslinking (see step 4). Tip: UV-crosslinking generally gives better results and enhanced sensitivity compared with baking. However, effective crosslinking requires optimization of the system.

Use either this step or step 4: To fix the DNA to the membrane by baking, first let the blot air-dry on a sheet of filter paper, then place between two sheets of filter paper, and bake at 80°C for 2 h. Proceed to step 5.

Use either this step or step 3: To fix the DNA by UV-crosslinking, first protect the surface of the membrane by covering the UV source (e.g., a transilluminator) with plastic wrap. Then take the damp blot and expose the side with DNA to the UV source for a predetermined length of time. Proceed to step 5. Tip: It is important to optimize the system for UV-crosslinking. To do this, prepare a blot with several control DNA samples. Cut the blot into separate strips for each lane, and irradiate each blot for different times, varying from 0.5 to 5 min. Hybridize all the blots together and determine which time gives the optimal signal intensity. It is important to use the same conditions (UV wavelength, distance from UV source) for each experiment. It is also important to calibrate the system routinely, as the energy emitted from a UV bulb is reduced with use. Tip: UV light can damage the eyes and skin. Always wear suitable eye and face protection.

If the blot will not be used immediately, store it at room temperature covered in plastic wrap.

Cleanup of DNA is often a prerequisite for efficient downstream applications such as cloning, sequencing, microarray analysis, or amplification.

The use of a dedicated kit for this application may be necessary, since kits can vary depending on the type of reaction and DNA fragment size (e.g., PCR products, gel extraction, enzymatic reactions, nucleotide removal, dye terminator removal) and the required elution volume.

Standard PCR cleanup only requires the removal of ~20 b oligos. In next-generation sequencing (NGS) library preparation, often, much larger primers — almost in the range of a PCR amplicon — must be removed, and PCR cleanup may not be sufficient. For this specialized “size selection” procedures, dedicated kits or PEG-based precipitation (8) are necessary.

This section describes considerations for isolation and quantification of RNA from different sample sources and RNA storage. It also deals with RNAi and the use of siRNA, together with miRNA, mimics, and inhibitors.

Whole genome amplification was developed as a way of increasing the amount of limited DNA samples. This is particularly useful for forensics and genetic disease research, where DNA quantities are limited, but many analyses are required. Various WGA techniques have been developed that differ both in their protocols, amplification accuracy, and ease-of-use.

Following the completion of the human genome project, the high demand for low-cost sequencing has given rise to a number of high-throughput, next-generation sequencing (NGS) technologies. These new sequencing platforms allow high-throughput sequencing for a wide range of applications.

The study of epigenetic mechanisms and DNA methylation has become increasingly important in many areas of research, including DNA repair, cell cycle control, developmental biology, cancer research, identification of biomarkers, predisposition factors, and potential drug targets.